An ex vivo Model of Paired Cultured Hippocampal Neurons for Bi-directionally Studying Synaptic Transmission and Plasticity

Synapses provide the main route of signal transduction within neuronal networks. Many factors regulate critical synaptic functions. These include presynaptic calcium channels, triggering neurotransmitter release, and postsynaptic ionotropic receptors, mediating excitatory and inhibitory postsynaptic potentials. The key features of synaptic transmission and plasticity can be studied in primary cultured hippocampal neurons. Here, we describe a protocol for the preparation and electrophysiological analysis of paired hippocampal neurons. This model system allows the selective genetic manipulation of one neuron in a simple neuronal network formed by only two hippocampal neurons. Bi-directionally analyzing synaptic transmission and short-term synaptic plasticity allows the analysis of both pre- and postsynaptic effects on synaptic transmission. For example, with one single paired network synaptic responses induced by both, a wild-type neuron and a genetically modified neuron can be directly compared. Ultimately, this protocol allows experimental modulation and hence investigation of synaptic mechanisms and thereby improves previously developed methods of studying synaptic transmission and plasticity in ex vivo cultured neurons. Key features Preparation of ex vivo paired cultured hippocampal neurons. Bi-directional electrophysiological recordings of synaptic transmission and plasticity. Genetic modulation of synaptic network formation (demonstrated by presynaptic viral overexpression of the auxiliary calcium channel α2δ-2 subunit). Graphical overview

Note: Paraffin can be transferred from PTFE dish to coverslips using fire-polished Pasteur pipettes ( Figure 1). Dip the pipette into paraffin and touch the coverslip quickly before paraffin hardens. Repeat procedure for each paraffin dot.   Bring the total volume to 4.5 mL with HBSS and add 0.5 mL of 2.5% trypsin (10 ×). Incubate in a water bath at 37 °C for 15 min. e. Remove the trypsin solution, add 5 mL of HBSS (gently tap or swirl the tube to mix), and let stand for 5 min. Repeat this step twice, finally bringing the volume to 3 mL (4 mL if hippocampi from more than five brains are used). f. Dissociate the cells by gently pipetting hippocampi up and down, first with a Pasteur pipette with a fire-polished tip to half the normal diameter, and then with a Pasteur pipette with a tip fire-polished to nearly a quarter the normal diameter. Continue pipetting gently until no chunks of tissue remain (approximately 7-8 times). g. Determine the density of cells using a hemacytometer. h. Add 25,000 cells to each of the dishes containing the poly-L-lysine-treated coverslips in neuronal plating medium (corresponding plating density is 880 cells/cm 2 ). i. For viral infection, add medium containing lentiviral particles to the dish with freshly added neurons.
Note: To reach an approximate 50% viral infection efficiency, the volume and concentration of lentivirus added should be defined experimentally. Added medium with lentiviral particles should not exceed 1 mL per Petri dish containing 4 mL of neuronal plating medium. j. After 3 h and using forceps, transfer the coverslips with the neurons attached into dishes containing the glial cells in neuronal maintenance medium (see step A3). Turn the coverslips upside down so that the neurons are facing down, towards the glial cells. k. To reduce glial proliferation, add Ara-C (5 μM) three days after plating the neurons. l. Once a week, remove 2 mL of the neuronal maintenance medium and replace it with fresh medium. m. For electrophysiological recordings, use cultured hippocampal neurons at the age 14-17 days in vitro (DIV) (Figure 2).

Figure 2. A pair of synaptically connected hippocampal neurons (left panel, phase contrast micrograph).
One neuron is virally transfected with the α2δ-2 protein and soluble eGFP (middle, fluorescence micrograph and right panel, overlayed phase contrast and fluorescent micrographs), the other neuron is an untransfected control neuron (arrow). Scale bar, 20 μm.

Preparation of astroglia feeder layer
Critical: It is necessary to start preparing the glia feeder layer 14 days prior to the preparation of hippocampal cultures. a. Prepare brain hemispheres as described in steps A2a-A2c.

Notes: i. If neuronal cultures are prepared on a regular basis, cells for the glia feeder layer can be prepared from the brains of mice used for the preparation of the neuronal culture.
ii. For all steps following the preparation of brain hemispheres, all media and reagents should be prewarmed to 37 °C before use. b. Mince brain hemispheres into small pieces with Vannas-Tübingen spring scissors. c. Transfer minced tissue to a 50 mL Falcon centrifuge tube in a final volume of 12 mL of HBSS. d. Add 1.5 mL of 1% DNase solution and incubate in water bath for 5 min at 37 °C. e. Add 1.5 mL of 2.5% trypsin (10×) and incubate for 15 min at 37 °C. During trypsin treatment, dissociate tissue every 5 min with a 5 mL serological pipette (pipette tissue up and down 7 -8 times). Critical: It is critical to add the DNase solution 5 min before adding the trypsin. Otherwise, DNase will be quickly degraded, resulting in abundant DNA material from minced brain tissues, which will strongly decrease the cell yield. f. Add 3 mL of horse serum to inhibit trypsin activity. g. Filter the combined supernatants through a 72 μm nylon mesh to remove any undissociated tissue. h. Centrifuge supernatants at 200× g for 5 min at 4 °C and resuspend cells in 5 mL of glial medium. i. Determine cell density with an hemacytometer. j. Transfer 4,000,000 cells into a 75 cm 2 T-flask (equivalent to approximately 1-1.5 brain hemispheres per flask). Add glial medium to a total volume of 13 mL. Put the flask in a cell culture incubator (37 °C, 5% CO 2 ). Note: One 70%-80% confluent T-flask with astroglia cells will be enough to prepare 10 mm × 60 mm dishes of astroglia feeding layer. k. Three days after plating, replace glial medium in the flask completely with fresh medium. l. Seven days after plating, shake the flask vigorously to dislodge microglia and remove them by washing (replace glia medium completely with fresh medium). Return the flask to the cell culture incubator.
Critical: Slap the flask forcefully 2-3 times on a hard surface so that the medium foams up and the entire content appears messed up. Gently tapping the flask will not dislodge the microglia, which are typically accumulating on top of astrocytes.
m. When cells in the flask have reached confluence (usually 10 days after plating), remove the glial medium and wash with 10 mL of HBSS. n. Remove HBSS, add 10 mL of 0.5% trypsin-EDTA (in HBSS), and put flask in cell culture incubator for 5 min. o. Tap the flask gently on the side to detached remaining astrocytes, add 1 mL of horse serum, and place the flask upright to allow the cells to slide to the bottom. p. Transfer glial cell suspension into a 50 mL Falcon centrifuge tube and centrifuge at 200× g for 5 min at room temperature. q. Resuspend cell pellet in 40 mL of glial medium. r. Add 4 mL of glial cell suspension per 60 mm Primaria plastic Petri dish. Put dishes in cell culture incubator (37 °C, 5% CO2). s. On the next day, replace glia media in Petri dishes with fresh medium. t. On the third day (one day before preparing hippocampal cultures), replace glia medium in dish es with astroglia layer with 6 mL of neuronal maintenance medium. Return dishes to the cell culture incubator.

B. Electrophysiological recordings of induced postsynaptic responses in paired cultured neurons
Notes: i. The following steps require the experimenter to have knowledge and proficiency in two -channel patch clamp electrophysiology. All recordings and analyses were performed using PatchMaster software.
ii. To induce synaptic transmission within paired neurons, each neuron will be interchangeably stimulated with a depolarization using an action potential wave form, which needs to be recorded from wild-type (WT) neurons.  c. Using probe 1, patch any neuron in voltage clamp mode using the whole-cell configuration. Holding potential (Vm) should be set to -70 mV.  f. In the Replay window, choose the recorded trace and zoom in on one action potential. g. Export the recorded action potential as a stimulation template file ( Figure 5).

General notes and troubleshooting
While performing experiments according to the current protocol, the experimenter may experience two types of problems related to the formation of neuronal pairs and the efficiency of viral infection. Both problems can be easily solved. If more than two neurons form networks on most of the poly-L-lysine spots (Figure 11), the plating density should be reduced (see step A2h). Increasing plating density is required if the majority of poly-L-lysine spots contain only one neuron. To successfully perform experiments on paired neurons in which one neuron was genetically modified, the efficiency of viral infection should be approximately 50%. If both neurons are virally infected or none are infected, it is necessary to decrease or increase, respectively, the concentration of the virus that is added to the freshly plated hippocampal neurons (step A2i).

Validation of protocol
The presented protocol was developed and successfully employed for the research work published in Geisler et al.
(2019), in order to analyze the consequences of altered synaptic wiring induced by the α2δ-2_ΔE23 isoform (section "Reduced synaptic transmission in aberrantly wired synapses," Figure 12 in the article).